Isolation, biochemical characterization, and genome sequencing of two high‐quality genomes of a novel chitinolytic Jeongeupia species

Abstract Chitin is the second most abundant polysaccharide worldwide as part of arthropods' exoskeletons and fungal cell walls. Low concentrations in soils and sediments indicate rapid decomposition through chitinolytic organisms in terrestrial and aquatic ecosystems. The enacting enzymes, so‐called chitinases, and their products, chitooligosaccharides, exhibit promising characteristics with applications ranging from crop protection to cosmetics, medical, textile, and wastewater industries. Exploring novel chitinolytic organisms is crucial to expand the enzymatical toolkit for biotechnological chitin utilization and to deepen our understanding of diverse catalytic mechanisms. In this study, we present two long‐read sequencing‐based genomes of highly similar Jeongeupia species, which have been screened, isolated, and biochemically characterized from chitin‐amended soil samples. Through metabolic characterization, whole‐genome alignments, and phylogenetic analysis, we could demonstrate how the investigated strains differ from the taxonomically closest strain Jeongeupia naejangsanensis BIO‐TAS4‐2T (DSM 24253). In silico analysis and sequence alignment revealed a multitude of highly conserved chitinolytic enzymes in the investigated Jeongeupia genomes. Based on these results, we suggest that the two strains represent a novel species within the genus of Jeongeupia, which may be useful for environmentally friendly N‐acetylglucosamine production from crustacean shell or fungal biomass waste or as a crop protection agent.

Despite its ubiquity, no significant long-term accumulation could be quantified in environmental soil or sediments, implying high turnover rates by chitinolytic organisms in nature (Gooday, 1990).
Most bacterial chitinases are classified as glycoside hydrolases of family 18 (GH18) and to a vastly lesser extent, those of family 19 (Cantarel et al., 2009). Chitin composition varieties in terrestrial and aquatic environments are reflected in the formation of distinct chitinolytic systems (Bai et al., 2016).
Aquatic chitinolytic bacteria might operate with a smaller toolkit of enzymes on average (Bai et al., 2016), are not enriched on the substrate (Brzezinska et al., 2008), and exhibit generally weaker catalytic activities (Swiontek Brzezinska et al., 2014). By contrast, terrestrial bacteria are more chitinolytically active in comparison, with Streptomyces as the predominant genus in the early stages of chitin decomposition, whereas other Actinomycetes take over the reins in later stages (de Boer et al., 1999;Swiontek Brzezinska et al., 2014).
Furthermore, a correlation between the abundance of bacteria and chitin decomposition rates could be observed in soil systems (Kielak et al., 2013), both of which could be promoted through the addition of substrate (Jacquiod et al., 2013;Mitchell & Alexander, 1962).
By virtue of their industrial potential and the increased relevance of sustainable (bio)technologies, extensive research on chitinases has been conducted (Binod et al., 2005;Juarez-Jimenez et al., 2008;Lan et al., 2004;Songsiriritthigul et al., 2010;Sun et al., 2019;Vaikuntapu et al., 2016). With 10% of the global crop loss arising from plant pathogens (Strange & Scott, 2005), chitinases could gain importance as environmentally friendly crop protection agents, in particular, due to their fungal cell walldirected hydrolase activities (Adrangi & Faramarzi, 2013;Gomaa, 2012;Neeraja et al., 2010;Veliz et al., 2017). However, turnover rates of recalcitrant chitin represent the biggest obstacle that hinders chitinases from becoming economically feasible contenders for industrial valorization. Thus, the exploration of novel chitinolytic organisms is important to further deepen our understanding regarding catalytic mechanisms and inferred optimization of enzymes. In this respect, recent improvements regarding the costs and accessibility of next-generation sequencing technologies enable the continuous democratization of wholegenome sequencing. Long-read sequencing platforms are well suited for de novo genome assembly applications, while highaccuracy short-read sequencing is apt for clinical variant discovery (Koboldt et al., 2013;Goodwin et al., 2016).
In this study, colloidal chitin amended soil samples were screened for chitinolytic organisms, isolated on chitin agar plates, and identified with 16S ribosomal RNA (rRNA) gene analysis (Jacquiod et al., 2013;Mitchell & Alexander, 1962 (Turrini et al., 2021) was utilized as a basis for taxonomic discussion. In addition, biochemical sugar metabolism capabilities were investigated utilizing API NE20 and CH50 stripes, revealing differences between the two strains investigated in this study and the type strain BIO-TAS4-2 T (Yoon et al., 2010).
Finally, in silico analysis of the chitinolytic systems demonstrated the highly conserved nature within the genus Jeongeupia and shed light on the enzymatic composition.

| Chemicals and consumables
All chemicals were supplied from Sigma-Aldrich, and general consumables were obtained from VWR. All necessary buffers and enzymes for next-generation genome sequencing were shipped from Pacific Biosciences. High molecular weight DNA was extracted with the Quick-DNA™ High Molecular Weight (HMW) MagBead Kit from Zymo Research and HMW genomic DNA (gDNA) shearing was conducted with g-TUBEs (Covaris) according to the manufacturer's manual.

| Colloidal chitin and media preparation
Colloidal chitin (CC) was prepared according to (Murthy & Bleakley, 2012) with slight modifications. Twenty grams of crab shell chitin powder (Sigma-Aldrich) were incrementally added to 150 mL 37% HCl under moderate stirring, increasing the viscosity of the solution. When the viscosity decreased sufficiently, more chitin was carefully added. The slur was then incubated for 2-3 h at room temperature under moderate stirring, evading the formation of bubbles. Afterward, the nonviscous, fully dissolved chitin of intense brown color was slowly poured into 2 L of ice-cold diH 2 O in a 5 L glass beaker and vigorously stirred, rapidly swelling to white colloidal chitin. The solution was incubated overnight at 4°C without stirring and neutralized the following day by adding excessive amounts of deionized water and subsequent centrifugation in a Beckman JLA8.1000 rotor for 15 min at 10,000g until pH 5 of the supernatant was reached. CC was harvested, autoclaved, and kept in the fridge until utilization in liquid chitinase screening media (CSM) or agar plates. The recipe was adapted and modified from (Lee et al., 1997;Singh et al., 1998): 20 g/L (2% wt/vol) CC, 0.7 g/L K 2 HPO 4 , 0.3 g/L KH 2 PO 4 , 0.5 g/L MgSO 4 × 5 H 2 O, 10 mg/L FeSO 4 × 7H 2 O, 20 g/L agar (optional), adjust to pH 6.5 for plates or 7 for liquid medium.
After autoclaving, 0.001 g/L ZnSO 4 and MnCl 2 were added from sterile filtrated stock solutions before pouring of agar plates/ inoculation of liquid media.

| Soil screening and cultivation of chitinolytic organisms
Soil samples were collected in sterile 50 mL falcon tubes and normalized to 60 g before transfer into 250 mL glass beakers. Tap water was added if the collected soil was completely dry. Afterward, samples were amended with either 1% or 10% wt/wt colloidal chitin or crab shell chitin powder (Sigma-Aldrich) and covered with tin foil.
After incubation at room temperature for 2 weeks, portions of the amended soil samples were transferred to sterile 50 mL falcon tubes and filled to 50 mL with sterile 1X PBS. Soil samples were incubated in a thermal shaker at 30°C and 600 rpm for 30 min. Supernatants were streaked out on CSM agar plates with different pH (5.5, 6, 6.5, 7, 8) using inoculation loops and incubated at 28°C for 2-3 days.
Colony-forming units (CFU) surrounded by halos were streaked onto separate CSM agar plates of the respective pH until axenic strains were obtained (Figure 1).

| Bacterial strains
Through the method described above, the two chitinolytic bacteria J. n. and J. sp. were isolated from environmental soil samples.

| Sequencing
Whole genome sequencing was performed on a Sequel IIe platform (Pacific Biosciences) on a single SMRT cell (lot number 418096) with the following parameters: 2 h of pre-extension, 2 h of adaptive loading (target p1 + p2 = 0.95) for a final on-plate concentration of 85 pM and a 30-h long movie window for signal detection (Ritz et al., 2023).
2. Genome quality assessment with CheckM (Parks et al., 2015) and Whole genome alignment was realized with the progressive-Mauve plugin within the Geneious Prime software v.2022.0.1, which is suitable for genomes containing rearranged segments due to recombination (Darling et al., 2010). Several locally collinear block (LCB) sizes were tested, whereby a compromise of conserved region count and sequence identity was selected, see Supporting Information: Data ( Figure A6).

| Phylogenetic trees with Type (Strain) Genome Server (TYGS)
The genome sequence data were uploaded to the TYGS, a free bioinformatics platform available at https://tygs.dsmz.de, for a The results were provided by the TYGS on 2023-05-16. The TYGS analysis was subdivided into the following steps: 1. Determination of closest type strain genomes: Was done in two complementary ways: First, all user genomes were compared against all type strain genomes available in the TYGS database via the MASH algorithm, a fast approximation of intergenomic relatedness (Ondov et al., 2016), and, the 10 type strains with the smallest MASH distances chosen per user genome. Second, an additional set of 10 closely related type strains was determined via the 16S rRNA gene sequences. These were extracted from the user genomes using RNAmmer (Lagesen et al., 2007). Each sequence was subsequently BLASTed (Camacho et al., 2009) against the 16S rRNA gene sequence of all 18,977-type strains currently available in the TYGS database. This was used as a proxy to find the best 50 matching type strains (according to the bitscore) for each user genome and to subsequently calculate precise distances using the Genome BLAST Distance Phylogeny approach (GBDP) under the algorithm "coverage" and distance formula d5 (Camacho et al., 2009;Meier-Kolthoff et al., 2013).
These distances were finally used to determine the 10 closest type strain genomes for each of the user genomes. 3. Phylogenetic inference: The resulting intergenomic distances were used to infer a balanced minimum evolution tree with branch support via FASTME 2.1.6.1 including subtree-prune-regraft moves postprocessing (Lefort et al., 2015). Branch support was inferred from 100 pseudobootstrap replicates each. The trees were rooted at the midpoint (Farris, 1972) and visualized with PhyD3 (Kreft et al., 2017).

| RESULTS AND DISCUSSION
3.1 | Screening, isolation, and 16S rRNA gene-based identification of chitinolytic bacteria Through the amendment of environmental soil samples with chitin in colloidal or powder form and dosages of 0.6% or 6% (wt/wt), respectively, chitinolytic microorganisms could putatively be enriched, as previously reported (Jacquiod et al., 2013). Streaking onto minimal media agar plates with 2% (wt/vol) colloidal chitin as the sole carbon-and nitrogen source produced CFUs, whose chitin hydrolyzing ability became visible through halos in varying diameters, indicating degradation of the paste-like, white colloidal chitin ( Figure A1). The two most promising candidates would then be subjected to 16S rRNA gene PCR ( Figure 1) and identified based on BLASTn comparison with the type strain database of NCBI (Sayers et al., 2022).
Both candidates were identified as J. naejangsanensis strain BIO-TAS4-2 T with identical percent identities of 98.72%, query coverages of 99%, and E values of 0. With identities of 98.48% (J. n.) and 98.01% (J. sp.), respectively, Jeongeupia chitinilytica's 16S rRNA gene showed the second most sequence homology. The transitory names were awarded based on these results, indicating that the investigated organism is J. naejangsanensis, leading to "J.

| Sugar metabolism
Carbon source utilization capabilities of the investigated strains were assessed using API 50CH and 20NE stripes (bioMérieux) and compared to the taxonomically closest strain J. naejangsanensis BIO-TAS4.2 T ( Table 1). As expected for closely related species, most examined characteristics were congruent, among these positive results for motility, nitrate reduction, N-acetylglucosamine, D-glucose, D-fructose, D-mannose, and D-ribose. Please refer to the Supporting Information: Data for a detailed list of all results and depictions of the API stripes.
Interestingly, certain differences could be illustrated regarding the assimilation of xylitol, D-lyxose, L-arabitol, and capric acid, all of which the type strain can utilize as a carbon source (Yoon et al., 2010).
Hydrolysis of the substrates esculin and gelatine was exclusive to J. n.
F I G U R E 1 Single streaks of the chitinolytic soil bacteria strains "J. n." (a) and "J. sp." (b) on colloidal chitin containing (2% wt/vol) agar plates. Strains were incubated at 28°C for 3 days before documentation. Chitinase screening media of different pH values were tested, pH 6 (a) and pH 7 (b) are depicted in this figure. Enzyme activity can be deduced by translucent halos around the colony-forming units, where chitin is degraded.
T A B L E 1 Phenotypic characteristics as determined with API 20NE and API 50CH stripes from bioMérieux.  and J. sp. on the other hand. In this regard, a minor metabolic distinction between the two strains described in this study could be made-with J. n. exhibiting a more potent esculin hydrolysis capability compared to J. sp., as detected with the API 20NE stripe ( Figure A4).  (Ciufo et al., 2018). This might also apply to the genus of Jeongeupia.  be accounted for when discussing genomic rearrangement or gene flux.

Hydrolysis of
Additionally, soil pH and salinity largely affect bacterial communities' composition (Fierer & Jackson, 2006;Lauber et al., 2009;Lozupone & Knight, 2007). Given the vast distance between the two sample collection sites, Naejang mountain in Korea and a billabong of the river Isar near Munich, Germany, the soil composition most likely differed. Furthermore, the distributed genome hypothesis, which states, that the gene pool of a bacterial taxon is more complex than that of an individual species, might serve to explain differences in observed genomes even within the species level, leading to genetic differences possibly reflected in ANI values (Baumdicker et al., 2012).  (Drula et al., 2021;Hemsworth et al., 2014Hemsworth et al., , 2015Henrissat et al., 2023;Mekasha et al., 2017;Slámová et al., 2010 (Kuusk et al., 2018;Westereng et al., 2017). Although the majority of research focused on fungal LPMOs (AA9 and AA11), their bacterial equivalents (AA10) are reported to boost the conventional hydrolytic activity of GH18 on chitin, as well (Forsberg et al., 2016;Vaaje-Kolstad et al., 2013). This way, 21 enzymes possibly involved in chitin degradation could be identified for J. n. and J. sp., respectively, comprising 13 GH18, 3 of which possess carbohydrate-binding modules of family 5, 3 GH19, 3 GH20, a single β-N-acetylhexosaminidase, and a single LPMO. Based on its published annotated draft genome (GCA_016865585.1), type strain J. naejangsanensis BIO-TAS4-2 exhibited 21 potentially chitinolytic enzymes, with 13 GH18, 3 GH19, 2 GH20, a single LPMO and one, as partial chitinase annotated putative protein (Turrini et al., 2021).
According to a study from 2016, which compared the chitinolytic systems of aquatic and terrestrial chitinolytic systems based on available genomes at that time, Jeongeupia exhibits an exceptionally rich enzyme toolkit (Bai et al., 2016), that reminds us of fungal Trichoderma species (Seidl et al., 2005). To our knowledge, few bacteria, among them Streptomyces coelicolor A3(2) (Saito et al., 2000) and Andreprevotia ripae (Lorentzen et al., 2021), are described with access to comparable chitinase gene copy numbers.
F I G U R E 3 Functional annotation of the J. n. and J. sp. genomes based on Clusters of Orthologous Groups (COGs) of proteins. Please note, that the scale for the group of poorly characterized enzymes (d) differs from that of the other functional groups (a-c). For the approx. A total of 3340 unique genes each, 79% could be annotated (a-c) while 21% are of unknown function (d).
To compare the chitinolytic systems taxonomically, and reveal orthologous enzymes, a CLUSTALW sequence alignment of the translated amino sequences was performed, followed by a phylogenetic tree generation (Huerta-Cepas, Serra, et al., 2016;Kyoto University Bioinformatics Center, 2023;Thompson et al., 1994) ( Figure 4). The results suggest that the chitinolytic enzymes of the three compared Jeongeupia strains are highly conserved, except for one orthologous GH20 unique to the two strains of this study and one single chitinase exclusive to the type strain reference genome.
Comprising the majority of bacterial chitinases, the GH18 were separated into three distinct clades, one of which could be functionally annotated as chitinase C by the eggNOG-mapper (Huerta-Cepas et al., 2017;Huerta-Cepas, Szklarczyk, et al., 2016).
The latter might represent the endo-chitinases, responsible for randomized cleavage along the chitin polysaccharide chain. Despite belonging to the same family, GH18 enzymes differentiate in sequence and catalytic mechanisms (Hoell et al., 2010), which is reflected by the two separate chitinase A-like branches, identified F I G U R E 4 Rootless phylogenetic tree of chitin-hydrolyzing Jeongeupia strains based on Shimodaira-Hasegawa-like local support. Enzymes were data mined from the Jeongeupia genomes with dbCAN 3.0, clades are labeled with Clusters of Orthologous Groups and Gene Ontology terms and SWISS-MODEL-based functional annotation predictions. Sequence alignment was performed with CLUSTALW. The phylogenetic tree was inferred using FastTree v2.1.8 with default parameters. BIO-TAS4-2 T = Jeongeupia naejangsanensis reference. J. n. and J. sp. are wholegenome sequenced strains from this study. Differences are framed in red. CBM, carbohydrate-binding module; GH, glycoside hydrolases of family 18, 19, or 20; LPMO, lytic polysaccharide monooxygenase.
with the SWISS-MODEL sequence homology database (Studer et al., 2020;Waterhouse et al., 2018). The auxiliary oxidoreductase enzyme LPMO was assigned to its own, distant branch based on sequence homology, and its oxygen-driven mechanism, which deviates drastically from conventionally operating hydrolase-based chitin-active enzymes (Bissaro et al., 2017;Kuusk et al., 2018).
Curiously, GH19 was represented in two separate clades, one seemingly homologous to a GH18 clade with carbohydrate-binding module 5, while the other clade shared more sequence identity with vastly different GH20 and β-N-acetyl-hexosaminidases.
All enzymes annotated as GH20 or β-N-acetyl-hexosaminidases, responsible for processive exo-chitinase activities, were assigned as descendants of a branch with three distinct clades. Since CLUSTALW is based purely on amino acid sequence alignment, the taxonomic allocation does not necessarily elucidate the singular clades' function but rather illuminates phylogenetic coherences and evolutionary processes.

| Comparison to the J. naejangsanensis
BIO-TAS4-2 T genome A whole-genome sequence alignment was conducted with the computational tool progressiveMauve (Darling et al., 2010) ( Figure 5). The software workflow includes selecting a reference sequence, followed by gapless multiple alignments of the input sequences, which serve as anchor regions. Subsequently, a phylogenetic guide tree is inferred, which is utilized to progressively apply an algorithm at every internal node, removing small matches that cause rearrangements and negatively affect the anchoring scores. Through an iterative process, progressiveMauve tries to align the sequences to maximize the conserved regions shared among the input sequences (Armstrong et al., 2019). PHASTER (Arndt et al., 2016;Zhou et al., 2011) revealed active and inactive phage regions, which are important driving forces of gene flux and microbial evolution (Canchaya et al., 2004;Mavrich & Hatfull, 2017). J. n. and J. sp. exhibit an identical phage region pattern, which might indicate that both strains are the same organism. On the other hand, the close relative J. naejangsanensis BIO-TAS4-2 T has fewer phage regions overall, with more inactive regions as depicted with gray in contrast to black stripes.
Circular plotting of the chitin-active gene loci elucidated different arrangements within the respective genomes. All chitin hydrolysis-related genes are clustered tightly in contig 1 of the reference genome, whereas the corresponding genes are distributed more evenly in the J. n./J. sp. genomes, with one GH18 in a particularly remote locus. Nevertheless, both chitinase C-like hydrolases in the genome reside in close proximity as well as two out of three GH19 and GH20 enzymes, respectively, forming small pseudo clusters.
Although the existence of distinct chitin hydrolase clusters might tempt one to assume varying enzymes, the alignment with CLUSTALW and the inferred phylogenetic tree depicted (Kyoto F I G U R E 6 Circos plot of the two in this study generated genome sequences of J. n. and J. sp., compared to the type strain genome of J. naejangsanensis BIO-TAS4-2 T . Circles from outermost to innermost represent (1) ideogram with contigs (ctg) and active phage regions indicated in black and inactive regions in gray, (2) conserved regions as detected with Mauve, (3) GC-skew; regions with above average GC contents are labeled in orange, in contrast to AT richer regions labeled in blue. The origin of replication (ORI) is usually located at one of the two transition points and was identified with DoriC. (4) Chitin-enacting enzyme CDS-accession numbers, due to clustering, not all proteins labels could be mapped, refer to Figure 4. Red = lytic polysaccharide monooxygenase, blue = N-acetyl-hexosaminidase, black = glycosyl hydrolase family 18 (GH18, chitinase), orange = GH19, green = GH20, (5) location of respective genes, and (6) links between homologous enzymes as identified with CLUSTALW amino acid sequence alignment. Image created with CIRCOS. *, this study; **, reference.
University Bioinformatics Center, 2023; Thompson et al., 1994) ( Figure 4), that the chitinolytic enzymes are highly conserved among the Jeongeupia genomes, but rearranged drastically. As suggested by the progressiveMAUVE alignment, most likely through gene flux events. Overall, the three genomes differ merely in two genes: J. naejangsanensis BIO-TAS4-2 T has an additional chitinase WP_239000134.1 which the strains of this study lack, whereas J.
n. and J. sp. have access to one additional GH20 hexosaminidase pgaptmp_000306/002118.
Lastly, GC-skew calculation could highlight over-and underabundance of the nucleotides guanine and cytosine. As a result, the two eligible ORI loci per genome could be unraveled, typically placed at the transition points of nucleotide overrepresentation (Lobry, 1996). Due to the replication initiation gene dnaA at one of those two conversion regions, the ORI could be located exactly with the DoriC 12.0 tool (Dong et al., 2023;Kosmidis et al., 2020;Trojanowski et al., 2018). Interestingly, the J. naejangsanensis
The novel Jeongeupia species presented in this study might provide a cost-effective and environmentally friendly process to convert crustacean shell and fungal biomass waste into Nacetylglucosamine based on its large set of chitin-active enzymes.
Further research must be conducted to demonstrate their suitability as antimycotic crop protection agents in a similar fashion to other studies (Neeraja et al., 2010;Swiontek Brzezinska et al., 2014). In addition, chitinases and other chitinoplastic enzymes such as chitindeacetylases could play significant roles in future circular bioeconomic approaches, where insects, crustaceans exoskeletons, or fungal residues are to be valorized in chemoenzymatic processes for applications in the food, chemical, cosmetic, and pharmaceutical industry (Intasian et al., 2021;Triunfo et al., 2022).

CONFLICT OF INTEREST STATEMENT
The authors declare no conflict of interest.

DATA AVAILABILITY STATEMENT
All data are provided in full in the results section and the appendix of this paper. All raw datasets generated and/or analyzed during the current study are available in the Zenodo online repository: https:// doi.org/10.5281/zenodo.8032359. The J. n. and J. sp. genomes can be accessed at NCBI with the BioSample IDs SAMN35557021 and SAMN35557022, respectively.

ETHICS STATEMENT
None required.

ORCID
Thomas B. Brück http://orcid.org/0000-0002-2113-6957 F I G U R E A1 Chitinase producer screening agar plates. Chitin-amended soil samples were incubated for 2 weeks at room temperature. The soil was moisturized with tap water if necessary. Then, sterile phosphate-buffered saline was added, and incubated for 30 min on a thermal shaker, and resulting supernatants were streaked out on chitin agar plates with pH 6 (a) and pH 7 (b). Halos around colony-forming units indicate chitin hydrolysis activities, due to degradation and therefore clearance of the white colloidal chitin. Colony forming units marked with an "X," highlighted through circles, were subsequently streaked on separate chitinase producer screening agar plates.
F I G U R E A2 API 50CH sugar metabolism results of the chitinolytic bacterium J. sp. (this study). Yellow highlights on the left indicate differences to the closely related J. naejangsanensis BIO-TAS4-2 type strain (Yoon et al., 2010). Test tubes colored in red indicate an inability to utilize a given sugar, while yellow test tubes indicate a positive result based on a pH shift. Esculin in tube 25 should turn black for a positive test result. We interpreted the strong darkening as a weak positive result. A1-A6 and Table A1.

See Figures
F I G U R E A3 API 50CH sugar metabolism results of the chitinolytic bacterium J. n. (this study). Yellow highlights on the left indicate differences to the closely related Jeongeupia naejangsanensis BIO-TAS4-2 type strain (Yoon et al., 2010). Test tubes colored in red indicate an inability to utilize a given sugar, and yellow test tubes indicate a positive result based on a pH shift. Esculin in tube 25 should turn black for a positive result. We interpreted the strong darkening as a weak positive result.
F I G U R E A5 Genome quality assessment with BUSCO (v.5.3.2), based on near-universal single-copy orthologs in the order of Neisseriales (odb10). Asterisk-labeled chitinolytic strains J. n. and J. sp. of this study were genome sequenced with a Sequel IIe platform (Pacific Biosciences). The reference genome Jeongeupia naejangsanensis BIO-TAS4-2 T was generated with a NextGen 500 platform (Illumina).